Sulfur Shuffle: Modulating Enzymatic Activity by Thiol-Disulfide

408

Bioconjugate Chem. 2000, 11, 408−413

Sulfur Shuffle: Modulating Enzymatic Activity by Thiol-Disulfide Interchange June M. Messmore,†,‡ Steven K. Holmgren,†,§ Juneko E. Grilley,† and Ronald T. Raines*,†,| Department of Biochemistry and Department of Chemistry, University of WisconsinsMadison, Madison, Wisconsin 53706. Received October 19, 1999; Revised Manuscript Received February 2, 2000

The facile modulation of biological processes is an important goal of biological chemists. Here, a general strategy is presented for controlling the catalytic activity of an enzyme. This strategy is demonstrated with ribonuclease A (RNase A), which catalyzes the cleavage of RNA. The side-chain amino group of Lys41 donates a hydrogen bond to a nonbridging oxygen in the transition state for RNA cleavage. Replacing Lys41 with a cysteine residue is known to decrease the value of kcat/Km by 105-fold. Forming a mixed disulfide between the side chain of Cys41 of K41C RNase A and cysteamine replaces the amino group and increases kcat/Km by 103-fold. This enzyme, which contains a mixed disulfide, is readily deactivated by dithiothreitol. Forming a mixed disulfide between the side chain of Cys41 and mercaptopropyl phosphate, which is designed to place a phosphoryl group in the active site, decreases activity by an additional 25-fold. This enzyme, which also contains a mixed disulfide, is reactivated in the presence of dithiothreitol and inorganic phosphate (which displaces the pendant phosphoryl group from the active site). An analogous control mechanism could be installed into the active site of virtually any enzyme by replacing an essential residue with a cysteine and elaborating the side chain of that cysteine into appropriate mixed disulfides.

INTRODUCTION

The activities of enzymes in vivo are exquisitely controlled by layers of regulatory mechanisms. These modalities are lost in vitro. Yet, the technological and commercial roles of enzymatic processes in vitro are of greater significance than ever before, with applications ranging from synthetic chemistry (Wong and Whitesides, 1994) to biotechnology (Eun, 1996) to food science (Suckling, 1990). Many practical uses of enzymes are likely to benefit from added degrees of control. Ribonucleases are enzymes that act at a crossroads of transcription and translation by catalyzing the cleavage of RNA (D’Alessio and Riordan, 1997). Controlling ribonucleolytic activity in vitro is especially important in manipulations in which the activity is required in some steps but detrimental in others. The ribonuclease protection assay (RPA),1 a method for detecting and quantitating RNA transcripts, is a prime example of this sort of manipulation (Zinn et al., 1983; Ausubel et al., 1994).2 * To whom correspondence should be addressed. Phone: (608) 262-8588. Fax: (608) 262-3453. E-mail: [email protected]. † Department of Biochemistry. ‡ Present address: Department of Biochemistry, Medical College of Wisconsin, Milwaukee, WI 53226. § Present address: Department of Chemistry and Biochemistry, Montana State UniversitysBozeman, Bozeman, MT 59717. | Department of Chemistry. 1 Abbreviations: CSA, cysteamine; DEAE, diethylaminoethyl; DTNB, 5,5′-dithiobis(2-nitrobenzoic acid); DTT, dithiothreitol; FAB, fast-atom bombardment; Hepes, N-(2-hydroxyethyl)piperazine-N′-2-ethanesulfonic acid; Mes, 2-(N-morpholino)-ethanesulfonic acid; MPP, 3-mercaptopropyl-1-phosphate; MS, mass spectrometry; NTB, 2-nitro-5-thiobenzoic acid; NMR, nuclear magnetic resonance; poly(C), poly(cytidylic acid); RNase A, bovine pancreatic ribonuclease A; RPA, ribonuclease protection assay; TEA, triethylamine; TLC, thin-layer chromatography; Tris, tris(hydroxymethyl)aminomethane.

Available methods for controlling ribonucleolytic activity tend to suffer from a lack of specificity, efficacy, or permanence. For example, diethyl pyrocarbonate effectively acylates the active-site histidine residues of ribonuclease A [RNase A; EC 3.1.27.5 (Raines, 1998)], destroying its enzymatic activity, but also covalently modifies nucleic acids and other proteins. Ribonucleoside complexes with vanadium(IV) and technetium(V) are rather weak inhibitors of catalysis by RNase A (Lindquist et al., 1973; Chen and Janda, 1992), and the vanadium(IV) complex is prone to air oxidation. In contrast, the complexes of RNase A with 3′,5′-pyrophosphate-linked deoxynucleotides have Kd values as low as 27 nM, but are likewise noncovalent (Leonidas et al., 1999; Russo and Shapiro, 1999). Although we and others have synthesized affinity labels and mechanism-based inactivators, the affinity labels bind weakly (Hummel et al., 1987) and the mechanismbased inactivators apparently modify nonessential residues (Stowell et al., 1995). Both types of compounds leave uninhibited a significant proportion of the ribonucleolytic activity. We have engineered a mechanism of in vitro control into RNase A. This method of modulating activity is similar to many natural regulatory mechanisms in that it employs reversible covalent modification. Specifically, 2 In an RPA, a labeled piece of anti-sense RNA is used to probe for an RNA transcript of interest. First, the labeled probe is added to a sample solution and allowed to hybridize with its cognate target RNA. Then, excess single-stranded probe is degraded by treatment with a ribonuclease, such as RNase A. Probe that has hybridized to target RNA is protected from degradation by the ribonuclease by virtue of being in a doublestranded complex with its cognate target RNA. The protected probe is then separated by electrophoresis in a polyacrylamide gel and visualized by autoradiography. If ribonucleolytic activity is not adequately controlled after the degradation step, both prior to and during electrophoresis, the signal resulting from the protected probe will be lost.

10.1021/bc990142m CCC: $19.00 © 2000 American Chemical Society Published on Web 04/09/2000

Modulating Enzymatic Activity by Thiol-Disulfide Interchange

we have replaced a Lys41 with a cysteine. Lys41 is known to donate a hydrogen bond to a nonbridging oxygen in the transition state for RNA cleavage (Messmore et al., 1995). The cysteine residue can be used as a “hook” on which to hang activating or deactivating entities. The K41C RNase A variant can be activated by chemical modification with cysteamine (2-aminoethanethiol), thus replacing the primary amine of the wild-type enzyme. Cysteamine can be removed later by gentle treatment with dithiothreitol (DTT) or another reducing agent. An inhibitory phosphoryl group at the end of a thiol-reactive linker can be added to achieve more complete inactivation. A few other enzymes (Pease et al., 1987; Gloss and Kirsch, 1995) and a lectin (Hollenbaugh et al., 1995) have been activated and deactivated by reversible modification of a newly introduced cysteine residue, but further inactivation via additional covalent changes to the cysteine residue is a feature unique to the present work. Similar regimens of transient activation and inactivation may be applicable to other enzymes with critical activesite lysine residues, including other nucleases and phosphotransferases. EXPERIMENTAL PROCEDURES

Materials. The oligonucleotide used for site-directed mutagenesis was synthesized bearing a trityl group and was purified using an Oligo Purification Cartridge from Applied Biosysytems (Foster City, CA). The Escherichia coli strain used for protein production, BL21(DE3), was from Novagen (Madison, WI). Ellman’s reagent, DTT, Hepes-free acid, Mes-free acid, and cysteamine‚HCl were from Sigma Chemical (St. Louis, MO). Poly(cytidylic acid) [poly(C)] was from either Midland Certified Reagents (Austin, TX) or Sigma Chemical. All starting materials for the synthesis of 2,4-dinitrophenyl 3-phosphoryl disulfide were from Aldrich Chemical (Milwaukee, WI), and the purity of these reagents was assessed by 1H NMR spectroscopy or thin-layer chromatography (TLC) (or both) prior to use. Spectroscopy and Protein Quantitation. Measurements of ultraviolet and visible absorption were made with a Cary model 3 spectrophotometer from Varian (Sugar Land, TX). Protein concentrations were determined based on the wild-type value of  ) 0.72 mL mg-1 cm-1 at 277.5 nm (Sela et al., 1957). This relationship was assumed to be valid for all forms of K41C RNase A studied herein. The veracity of this assumption was supported by DTT treatment of K41C-ntb RNase A (vide infra) yielding the expected value of 1.0 ( 0.2 equiv of 2-nitro-5-thiobenzoic acid (NTB). Preparation of K41C-(2-nitro-5-thiobenzoic acid) RNase A. A cDNA encoding K41C RNase A (Messmore et al., 1995) was expressed in E. coli strain BL21(DE3) under the control of the T7 RNA polymerase promoter. The resulting protein was purified as described previously (delCardayre´ et al., 1995; Messmore et al., 1995). After purification, the new sulfhydryl group at position 41 was protected from inadvertent reaction by treatment with 5,5′-dithiobis(2-nitrobenzoic acid), Ellman’s reagent or DTNB (Ellman, 1959). The protected protein, K41Cntb RNase A, was separated from any unprotected protein by cation exchange FPLC (MonoS column; Pharmacia, Piscatawy, NJ) using a linear gradient of NaCl (0 to 200 mM) in 50 mM Hepes-NaOH buffer (pH 7.7). The protected protein eluted from the column at 60 mM NaCl, and the unprotected K41C protein eluted at 80 mM. The protected protein was stable for several months when stored at 4 °C.

Bioconjugate Chem., Vol. 11, No. 3, 2000 409

Preparation of K41C-Cysteamine RNase A. The thiol-disulfide interchange reaction between K41C-ntb RNase A and cysteamine (CSA) was initiated by adding cysteamine‚HCl (2-10 µL of a 0.010 M solution in 0.10 M Tris-HCl buffer, pH 7.7) to a solution (1.0 mL) of 50 mM Hepes-NaOH buffer (pH 7.7) containing K41C-ntb RNase A (0.3-0.7 mg/mL). The reaction mixture was incubated at 25 °C, and its progress was monitored by following the increase in absorbance at 412 nm, resulting from the release of a stoichiometric quantity of NTB [ ) 13 600 M-1 cm-1 (Ellman, 1959)]. Excess cysteamine and released NTB were removed by subsequent dialysis. Assay of Poly(cytidylic acid) Cleavage. Ribonucleolytic activity was assessed by using poly(C) as a substrate. Prior to an assay, poly(C) was precipitated from aqueous ethanol (70% v/v), solubilized in assay buffer, and quantitated for total cytidyl concentration by absorbance at 268 nm using  ) 6200 M-1 cm-1 (Yakovlev et al., 1992). Assays of poly(C) cleavage were performed at 25 °C in 0.10 M MES-NaOH buffer (pH 6.0) containing NaCl (0.10 M). Cleavage of poly(C) was monitored by the increase in absorbance at 250 nm using ∆ ) 2380 M-1 cm-1 (delCardayre´ and Raines, 1994). Concentrations of poly(C) in assays ranged from 0.02 to 1.2 mM, in terms of individual cytidyl units. Initial rates of cleavage were fitted to the Michaelis-Menten equation with the program HYPERO (Cleland, 1979). Deactivation of K41C-Cysteamine RNase A with Dithiothreitol. The reduction of the mixed disulfide in K41C-csa RNase A (20 µM) by DTT (20 µM or 0.32 mM) was performed in 50 mM Hepes-NaOH buffer (pH 7.7). The reaction was allowed to proceed at 25 °C, and aliquots were removed periodically to assay for ribonucleolytic activity. Synthesis of 2,4-Dinitrophenyl 3-Phosphopropyl Disulfide (7). Disulfide 7 was synthesized by the route shown in Scheme 1. p-Methoxybenzylmercaptan (1). p-Methoxybenzylmercaptan was prepared from p-methoxybenzyl chloride and thiourea. A 1:1 ratio of the reactants was boiled under reflux and N2(g) in acetonitrile for 1 h, and then stirred at room temperature. After 12 h, the reaction mixture was concentrated under reduced pressure to give a white solid. To hydrolyze the amidine groups, the solid was then dissolved in an aqueous solution of potassium carbonate and sodium bisulfite, and this solution was heated to 80 °C. The solution was subsequently acidified, and the product was extracted into chloroform, dried over MgSO4, and evaporated to give a slightly yellow oil. 1H NMR (CDCl3, ppm): 7.22-7.25 and 6.86-6.83 (two m, 2H each, -C6H4-), 3.79 (s, 3H, -OCH3), 3.70 (d, J ) 7.4 Hz, 2H, -CH2SH), 1.73 (t, J ) 7.4 Hz, 1H, -SH). 3-Hydroxypropyl p-Methoxybenzyl Thioether (2). Equimolar amounts of p-methoxybenzylmercaptan and 3-bromo1-propanol in acetonitrile were stirred under N2(g) in the presence of potassium carbonate. The reaction was allowed to proceed, with stirring, overnight at room temperature. Compound 3 was isolated by silica chromatography, with 1:1 ethyl acetate:hexane as the mobile phase, and then evaporated under reduced pressure to give a colorless oil. 1H NMR (CDCl3, ppm): 7.21-7.26 and 6.846.87 (two m, 2H each, MeOC6H4-), 3.80 (s, 3H, -OCH3), 3.71 (t, J ) 6.1 Hz, 2H, -CH2OH), 3.68 (s, 2H, -C6H4CH2S-), 2.53 (t, J ) 7.0 Hz, 2H, -SCH2CH2-), 1.81 (m, 2H, -CH2CH2CH2-), 1.61 (br s, 1H, CH2OH). 3-Phosphopropyl p-Methoxybenzyl Thioether (6). Compound 6 was synthesized by a method analogous to that of (Kraszewski and Stawinski, 1980). Bis-triazole pnitrophenyl phosphate (3) was generated by the reaction

410 Bioconjugate Chem., Vol. 11, No. 3, 2000

Messmore et al.

Scheme 1

of p-nitrophenyl-phosphorodichloridate plus triazole and triethylamine (TEA) in tetrahydrofuran under N2(g), first at 0 °C and then at room temperature, for a total of 30 min. Compound 2 (500 mg; 2.36 mmol) was then added in a 1:3 molar ratio relative to the dichloridate, along with 1.3 equiv of 1-methylimidazole relative to 3. The reaction was allowed to proceed, still under N2(g), for another 30 min. H2O and TEA were then added to hydrolyze the triazole groups. After 2 h, the reaction mixture was concentrated under reduced pressure to give a yellow solid. The yellow solid was dissolved in chloroform and washed with aqueous NaHCO3, then concentrated to give a yellow oil. The yellow oil was dissolved in H2O and purified by two chromatographic steps, ion exchange (Sephadex SP C-25) and RP-C18 (isocratic; 1:1 MeOH: H2O mobile phase), to give the phosphodiester 5 in 62% yield from alcohol 2. MS of 5 (FAB, m/e): 412.1 (calcd for C17H19NO7PS, 412.06). Phosphodiester 5 was hydrolyzed in 1 N NaOH at 90 °C for 16 h, and the resulting phosphomonoester (6) was neutralized and purified by anion-exchange chromatographically on DEAE Sephadex resin. Elution from the resin was effected by a linear gradient (0.1 to 1.0 M) of TEA-H2CO3, which was removed as the product, was dried under reduced pressure, yielding 6 in 75% yield from 5. 1H NMR (D2O, ppm): 7.06-7.10 and 6.72-6.76 (two m, 2H each, MeOC6H4-), 3.73 (quartet, J ) 6.4 Hz, 2H, -CH2OPO3H2), 3.61 (s, 3H, -OCH3), 3.50 (s, 2H, -C6H4CH2S-), 2.37 (t, J ) 7.5 Hz, 2H, -SCH2CH2-), 1.70 (quintet, J ) 6.6 Hz, 2H, -CH2CH2CH2-). 31P NMR (D2O, ppm relative to 85% w/v phosphoric acid): 1.36 (s, decoupled). Dinitrophenyl 3-Phosphopropyl Disulfide (7). The simultaneous removal of the p-methoxybenzyl group from the sulfur and creation of the disulfide was accomplished by the method of (Johnson et al., 1994). Thioether 6 was stirred at room temperature for 30 min with an equimolar amount of 2,4-dinitrophenylsulfenyl chloride in acetic acid. The reaction mixture was then concentrated by

evaporation and dissolved in methanol. The product was crystallized with the addition of diethyl ether and isolated by filtration to yield disulfide 7 as the free acid in 35% yield from 6. 1H NMR (CD3OD, ppm): 9.04 and 8.588.62 [s and m, respectively, 1H and 2H, respectively, (NO2)2C6H3-], 4.06 (quartet, J ) 6.2 Hz, 2H, -CH2CH2OPO3H2), 2.96 (t, J ) 7.2 Hz, 2H, -SCH2CH2-), 2.05 (quintet, J ) 6.5 Hz, 2H, -CH2CH2CH2-). 31P NMR (D2O, ppm relative to phosphoric acid): 0.8 (s, decoupled). K41C-(3-Mercaptopropyl-1-phosphate) RNase A. K41C RNase A was isolated freshly from K41C-ntb RNase A by DTT treatment and cation-exchange chromatography on a mono S column (vide supra). The protein (at 6 µM) was then reacted with a 10-fold molar excess of 7, and then reisolated by cation-exchange chromatography. The addition of the MPP moiety caused the protein to elute from the Mono S column at 45 mM NaCl, rather than at 80 mM NaCl. Zymogram Assay. Extremely small amounts of ribonucleolytic activity can be detected in Laemmli gels impregnated with RNA or poly(C) (Blank et al., 1982; Kim and Raines, 1993; Bravo et al., 1994). Here, such zymogram assays were performed in an polyacrylamide (18% w/v) gel containing poly(C) (0.5 mg/mL). K41C RNase A (0.4 µg) or K41C-mpp RNase A (0.4 µg) were loaded onto the gel in a nonreducing glycerol loading buffer (105-fold greater

than the actual molarity of the phosphoryl group of MPP in our assays ([K41C-mpp RNase A] e 3.5 µM). Once K41C RNase A has been modified with MPP, noncovalent interactions with the pendant phosphoryl group increased markedly the stability of the intermolecular disulfide bond. DTT and β-mercaptoethanol were used over a range of conditions that are benign to the K41C enzyme in attempts to remove the MPP group from K41C-mpp RNase A. At low and moderate ionic strengths, no increase in activity was observed by zymogram analysis after incubation with up to 2 mM DTT for up to 24 h. Even in the presence of 1 M inorganic phosphate and 0.10 mM DTT, no measurable gain in activity was observable. At 2.5 M inorganic phosphate and 2 mM DTT, a gain of activity indicative of a 20% reversal to K41C RNase A was observed after 90 min. Conclusion. The unique reactivity of the sulfhydryl group allows for rapid and specific control of enzymatic activity. The activity of K41C RNase A has been increased 1100-fold by modification with cysteamine, and decreased 25-fold by modification with 3-mercaptopropyl1-phosphate. Enzymatic activity can thus be readily modulated over a range spanning a factor of 104.4-fold. An analogous control mechanism could conceivably be installed into the active site of any enzyme if an essential functional group is available for replacement with a cysteine. ACKNOWLEDGMENT

We are grateful to C. L. Dantzman for advice on the synthesis of dinitrophenyl 3-phosphopropyl disulfide (7) and K. J. Woycechowsky and C. L. Jenkins for comments on the manuscript. This work was supported by Grants GM44783 (NIH) and BES 9604563 (NSF). S.K.H. was an Arthritis Foundation postdoctoral fellow. LITERATURE CITED Anderson, D. G., Hammes, G. C., and Walz, F. G. J. (1968) Binding of Phosphate Ligands to Ribonuclease A. Biochemistry 7, 1637-1645. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K., Eds. (1994) Current Protocols in Molecular Biology, John Wiley & Sons, Boston. Blank, A., Sugiyama, R. H., and Dekker, C. A. (1982) Activity staining of nucleolytic enzymes after SDS-PAGE: use of aqueous isopropanol to remove detergent from gels. Anal. Biochem. 120, 267-275. Bravo, J., Ferna´ndez, E., Ribo´, M., de Llorens, R., and Cuchillo, C. M. (1994) A versatile negative-staining ribonuclease zymogram. Anal. Biochem. 219, 82-86. Chen, Y.-C. J., and Janda, K. D. (1992) A new approach toward the inhibition of ribonucleases: a water-stable ribonucleosidetechnetium chelate. J. Am. Chem. Soc. 114, 1488-1489. Cleland, W. W. (1979) Statistical analysis of enzyme kinetic data. Methods Enzymol. 63, 103-138. Cleland, W. W., and Northrop, D. B. (1999) Energetics of substrate binding, catalysis, and product release. Methods Enzymol. 308, 3-27. D’Alessio, G., and Riordan, J. F., Eds. (1997) Ribonucleases: Structures and Functions, Academic Press, New York. delCardayre´, S. B., and Raines, R. T. (1994) Structural Determinants of Enzymatic Processivity. Biochemistry 33, 60326037. delCardayre´, S. B., Ribo´, M., Yokel, E. M., Quirk, D. J., Rutter, W. J., and Raines, R. T. (1995) Engineering ribonuclease A: production, purification, and characterization of wild-type enzyme and mutants at Gln11. Protein Eng. 8, 261-273. Ellman, G. L. (1959) Tissue sulfhydryl groups. Arch. Biochem. Biophys. 82, 70-77.

Modulating Enzymatic Activity by Thiol-Disulfide Interchange Eun, H.-M. (1996) Enzymology Primer for Recombinant DNA Technology, Academic Press, San Diego. Faulstich, H., and Heintz, D. (1995) Reversible introduction of thiol compounds into proteins by use of activated mixed disulfides. Methods Enzymol. 251, 357-366. Gloss, L. M., and Kirsch, J. F. (1995) Examining the structural and chemical flexibility of the active site base, Lys-258, of Eschericia coli aspartate aminotransferase by replacement with unnatural amino acids. Biochemistry 34, 12323-12332. Hollenbaugh, D., Aruffo, A., and Senter, P. D. (1995) Effects of Chemical Modification on the Binding Activities of P-Selectin Mutants. Biochemistry 34, 5678-5684. Hummel, C. F., Pincus, M. R., Brandt-Rauf, P. W., Frei, G. M., and Carty, R. P. (1987) Reaction of (Bromoacetamido)nucleoside Affinity Labels with Ribonuclease A: Evidence for Steric Control of Reaction Specificity and Alkylation Rate. Biochemistry 26, 135-146. Johnson, R., Joshi, B. V., and Reese, C. B. (1994) Synthesis of 2′,3′-dithiouridine. J. Chem. Soc., Chem. Commun., 133-134. Kim, J.-S., and Raines, R. T. (1993) Ribonuclease S-peptide as a carrier in fusion proteins. Protein Sci. 2, 248-356. Kraszewski, A., and Stawinski, J. (1980) Phosphoryl tristriazolesa new phosphorylating reagent. Tetrahedron Lett. 21, 2935-2936. Leonidas, D. D., Shapiro, R., Irons, L. I., Russo, N., and Acharya, K. R. (1999) Toward Rational Design of Ribonuclease Inhibitors: High-Resolution Crystal Strcuture of a Ribonuclease A Complex with a Potent 3′,5′-Pyrophosphate-Linked Dinucleotide Inhibitor. Biochemistry 38, 10287-10297. Li, Y.-J., Rothwarf, D. M., and Scheraga, H. A. (1995) Mechanism of reductive protein unfolding. Nat. Struct. Biol. 2, 489-494. Lindquist, R. N., Lynn, J. L., Jr., and Lienhard, G. E. (1973) Possible Transition-State Analogs for Ribonuclease. The Complexes of Uridine with Oxovanadium(IV) Ion and Vanadium(V) Ion. J. Am. Chem. Soc. 95, 8762-8768. Lundblad, R. L. (1991) Chemical Reagents for Protein Modification, CRC Press, Boca Raton, FL. Messmore, J. M., Fuchs, D. N., and Raines, R. T. (1995) Ribonuclease A: Revealing Structure-Function Relationships with Semisynthesis. J. Am. Chem. Soc. 117, 8057-8060.

Bioconjugate Chem., Vol. 11, No. 3, 2000 413 Pease, M. D., Serpersu, E. H., and Mildvan, A. S. (1987) Chemical modification and spin-labelling studies of the Arg 87 to Cys (R87C) mutant of Staphylococcal nuclease. Fed. Proc. 46, 1932. Raines, R. T. (1997) Nature’s transitory covalent bond. Nat. Struct. Biol. 4, 424-427. Raines, R. T. (1998) Ribonuclease A. Chem. Rev. 98, 10451065. Russo, N., and Shapiro, R. (1999) Potent inhibition of mammalian ribonucleases by 3′,5′-pyrophosphate-linked nucleotides. J. Biol. Chem. 274, 14902-14908. Sela, M., Anfinsen, C. B., and Harrington, W. F. (1957) The correlation of ribonuclease activity with specific aspects of tertiary structure. Biochim. Biophys. Acta 26, 502-512. Stowell, J. K., Widlanski, T. S., Kutateladze, T. G., and Raines, R. T. (1995) Mechanism-Based Inactivation of Ribonuclease A. J. Org. Chem. 60, 6930-6936. Suckling, C. J., Ed. (1990) Enzyme Chemistry: Impact and Applications, Chapman and Hall, London. Viola, R. E., and Wright, S. K. (1999) New enzymes from old. In Enzymatic Mechanisms (P. A. Frey and D. B. Northrop, Eds.) pp 116-130, IOS Press, Washington DC. Wlodawer, A. (1985) Structure of bovine pancreatic ribonuclease by X-ray and neutron diffraction. In Biological Macromolecules and Assemblies, Vol. II, Nucleic Acids and Interactive Proteins (F. A. Jurnak and A. McPherson, Eds.) pp 395-439, Wiley, New York. Wong, C.-H., and Whitesides, G. M. (1994) Enzymes in Synthetic Organic Chemistry, Elsevier Science, Tarrytown, NY. Yakovlev, G. I., Moiseyev, G. P., Bezborodova, S. I., Both, V., and Sevcik, J. (1992) A comparative study on the catalytic properies of guanylyl-specific ribonucleases. Eur. J. Biochem. 204, 187-190. Zinn, K., DiMaio, D., and Maniatis, T. (1983) Identification of two distinct regulatory regions adjacent to the human β-interferon gene. Cell 34, 865-879.

BC990142M